New Bio-Indicators for Long Term Natural …Compounds in Deep Terrestrial Aquifers. Front....

ORIGINAL RESEARCHpublished: 09 February 2016

doi: 10.3389/fmicb.2016.00122

Edited by:John Joseph Kilbane,

Illinois Institute of Technology/Intertek,USA

Reviewed by:Uwe Strotmann,

Westfaelische Hochschule, GermanyTapan Kumar Adhya,KIIT University, India

*Correspondence:Anthony Ranchou-Peyruse

[email protected]

†These authors are co-first authors.

Specialty section:This article was submitted to

Microbiotechnology, Ecotoxicologyand Bioremediation,

a section of the journalFrontiers in Microbiology

Received: 08 December 2015Accepted: 22 January 2016

Published: 09 February 2016

Citation:Aüllo T, Berlendis S, Lascourrèges

J-F, Dessort D, Duclerc D,Saint-Laurent S, Schraauwers B,

Mas J, Patriarche D, Boesinger C,Magot M, Ranchou-Peyruse A (2016)

New Bio-Indicators for Long TermNatural Attenuation of Monoaromatic

Compounds in Deep TerrestrialAquifers. Front. Microbiol. 7:122.doi: 10.3389/fmicb.2016.00122

New Bio-Indicators for Long TermNatural Attenuation of MonoaromaticCompounds in Deep TerrestrialAquifersThomas Aüllo1†, Sabrina Berlendis1†, Jean-François Lascourrèges2†, Daniel Dessort3,Dominique Duclerc3, Stéphanie Saint-Laurent1, Blandine Schraauwers2, Johan Mas1,Delphine Patriarche4, Cécile Boesinger5, Michel Magot1 andAnthony Ranchou-Peyruse1*

1 Université de Pau et des Pays de l’Adour, Institut des Sciences Analytiques et de Physico-Chimie Pour l’Environnement etles Matériaux UMR 5254, Equipe Environnement et Microbiologie, Pau, France, 2 APESA, Pau, France, 3 TOTAL –Centre-Scientifique-Technique-Jean-Feger, Pau, France, 4 STORENGY – Geosciences Department, Bois-Colombes, France,5 TIGF – Transport et Infrastructures Gaz France, Pau, France

Deep subsurface aquifers despite difficult access, represent important water resourcesand, at the same time, are key locations for subsurface engineering activities for theoil and gas industries, geothermal energy, and CO2 or energy storage. Formationwater originating from a 760 m-deep geological gas storage aquifer was sampledand microcosms were set up to test the biodegradation potential of BTEX byindigenous microorganisms. The microbial community diversity was studied usingmolecular approaches based on 16S rRNA genes. After a long incubation period,with several subcultures, a sulfate-reducing consortium composed of only twoDesulfotomaculum populations was observed able to degrade benzene, toluene, andethylbenzene, extending the number of hydrocarbonoclastic–related species among theDesulfotomaculum genus. Furthermore, we were able to couple specific carbon andhydrogen isotopic fractionation during benzene removal and the results obtained bydual compound specific isotope analysis (εC = −2.4 ± 0.3 ; εH = −57 ± 0.98 ;AKIEC: 1.0146 ± 0.0009, and AKIEH: 1.5184

� � � �± 0.0283) were close to those

obtained previously in sulfate-reducing conditions: this finding could confirm theexistence of a common enzymatic reaction involving sulfate-reducers to activatebenzene anaerobically. Although we cannot assign the role of each population ofDesulfotomaculum in the mono-aromatic hydrocarbon degradation, this study suggestsan important role of the genus Desulfotomaculum as potential biodegrader amongindigenous populations in subsurface habitats. This community represents the simplestmodel of benzene-degrading anaerobes originating from the deepest subterraneansettings ever described. As Desulfotomaculum species are often encountered insubsurface environments, this study provides some interesting results for assessingthe natural response of these specific hydrologic systems in response to BTEXcontamination during remediation projects.

Keywords: natural attenuation, deep aquifer, BTEX, sulfate-reduction, Desulfotomaculum

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Aüllo et al. Hydrocarbons Biodegradation in Deep Aquifers

INTRODUCTION

Deep subterranean ecosystems have been described duringthe last decades as a key living earth component for globalcarbon cycling and geo-engineering system (Pedersen, 2000;Griebler et al., 2014; Wilkins et al., 2014). This is explainedby the unexpected microbial biomass discovered leading to thepresumption that these ecosystems potentially host an estimatedbiomass equivalent to about 40–60% of the terrestrial surfacebiomass (Whitman et al., 1998; McMahon and Parnell, 2014).However, the available ecological data concerning subterraneanenvironments remain limited in literature, mainly because ofthe difficulties in collecting representative samples, especially indeep confined aquifers. Here, we imply deep confined aquifersto be geological formations located 100s of meters deep andisolated from surface interaction by an impermeable geologicallayer. These geological formations are often associated with activepetroleum reservoirs, or depleted oil fields used for undergroundgas storage.

The biodegradation of hydrocarbons has long been regardedas a strictly aerobic process, depending on oxygen availability andthe presence of oxygen-respiring bacteria. But during the last fewdecades, anaerobic hydrocarbon degradation has been describedin anaerobic surface or shallow subsurface environments (Barkeret al., 1987; Ball and Reinhard, 1996; Head et al., 2003, 2010;Widdel et al., 2010) by numerous consortia and several originalbacterial strains (Heider et al., 1999; Widdel and Rabus, 2001;Weelink et al., 2010; Kuppardt et al., 2014). It has been shown thatBTEX (Benzene, Toluene, Ethylbenzene, and Xylenes isomers)could be anaerobically degraded using a variety of terminalelectron acceptors such as sulfate, nitrate, ferric iron, and CO2(Anderson and Lovley, 2000; Chakraborty and Coates, 2004;Weelink et al., 2010; Holmes et al., 2011; Vogt et al., 2011).However, benzene and ethylbenzene are the most recalcitrantof these hydrocarbons. No pure strain able to degrade benzenein sulfate-reducing conditions has been isolated, and only oneregarding ethylbenzene (Kniemeyer et al., 2003).

Oil biodegradation in petroleum reservoirs, and by extensionin all deep subsurface environments, is an anaerobic process(Magot et al., 2000; Head et al., 2003; Widdel et al., 2010).Very little is known about bacterial species involved in similarprocesses in the terrestrial subsurfaces, despite the importanceof oil biodegradation to the oil industry. Some studies havereported direct or indirect evidences of oil biodegradation bymolecular ecology studies (Nazina et al., 2006), thermodynamiccalculations (Dolfing et al., 2008; Onstott et al., 2010), isotopicfractionation and 13C tracer-labeled experiments in microcosms(Mancini et al., 2003; Elsner et al., 2005; Fischer et al., 2008; Joneset al., 2008; Imfeld et al., 2014) and by cultural approaches underhigh temperature and pressure mimicking petroleum reservoirconditions (Mayumi et al., 2011; for a review, see Head et al.,2010).

Studying the microbial ecology and microbial activities ofthe deep subsurface is difficult, mainly because representativesamples are technically very challenging to collect in these deepenvironments and microbiological studies are scarce (Greksáket al., 1990; Ivanova et al., 2007; Balk et al., 2008; Balk et al.,

2010). The opportunity to collect representative fluids fromthe deep subsurface occurred for a few years with a specificsampling protocol set up for control wells of natural gas storagein deep aquifers (Basso et al., 2005, 2009). This led to the recentobservation that original anaerobic microbial communitiescollected from a 830 m deep gas storage aquifer were involved inthe natural attenuation of BTEX in these hydrocarbon-impactedenvironments (Berlendis et al., 2010). Different microbialconsortia were obtained depending on the culture conditions andhydrocarbons used as carbon sources. Proteobacteria, Firmicutesrelated to the Desulfotomaculum, Chlorobi, Thermotogales,Bacteroidetes, Synergistes, and Euryarchaeota were shown to bepresent in the BTEX-degrading consortia by 16S rRNA genestudies, but specific hydrocarbon degradation activity has notbeen linked to any specific bacterial or archaeal group. Microbialinventory investigations of potential biodegraders isolated fromdeep confined subterranean environments are still of crucialinterest for geo-engineering activities from oil productionto bioremediation strategies. Therefore, we investigated theselection of potential biodegraders from another deep gas-storagesubterranean aquifer. The microbial community strongly differsfrom that previously reported (Berlendis et al., 2010) and wasable to anaerobically degrade benzene, ethylbenzene, and tolueneduring 10 years of culturing. It confirms the presence of spore-forming bacteria belonging to the genus Desulfotomaculumas another set of autochthonous mono-aromatic hydrocarbonsbiodegraders in deep subsurface environments under sulfate-reducing conditions.

MATERIALS AND METHODS

SamplesFormation water was sampled from a deep subterranean aquifer(Parisian basin, France) located at 760 m of depth in a Jurassicsuperior geological formation (Lusitanien, calcareous oolites).This aquifer is confined in a poorly carbonated sandstoneformation by an overlaying impermeable geological layer. Theaquifer is used for geological storage of natural gas. In situtemperature and pH were 37◦C and 8.2, respectively. Thetotal salinity of water was 1.6 g.L−1. Formation water andconcentrated biomass (Sterivex Filter units, EMDMillipore) werecollected anoxically from the wellhead of a peripheral monitoringwell after a specific cleaning procedure as previously described(Basso et al., 2005). The samples filtered on-site used in thisstudy were transported to the laboratory under anoxic conditions(GasPakTM EZ, BD), stored at 4◦C to avoid microbial growth andprocessed the day after.

Microcosm ExperimentsThe concentrated microflora collected on site on 78 0.2-μm-pore-size SterivexTM filters (Millipore) were resuspended in2.6 L of anoxic formation water. The 102-fold concentratedbacterial suspension was used as inoculum in several flaskswith formation water supplemented with 0.5 g.L−1 NH4Cl,0.3 g.L−1 K2HPO4, 0.3 g.L−1 KH2PO4, and 2 g.L−1 Na2SO4for sulfate-reducing conditions, or 0.085 g.L−1 NaNO3 for






nitrate-reducing conditions, or 0.3 g.L−1 FeIII-citrate for iron-reducing conditions, or flushed with CO2/H2 (20/80) formethanogenic/fermentative conditions. For each condition, themedia were supplemented with 1 mL.L−1 of trace-elementssolution and 1 mL.L−1 of vitamin solution from a sterileanoxic stock solution prepared under N2 (Pfennig et al.,1981; Eichler and Pfennig, 1986). One milliliter per liter ofdithionite solution (0.2% w/v) was added to the media asa reducing agent and resazurine (1 mg.L−1) was used as aredox indicator. From all the initial conditions (sulfate-, nitrate-,iron-reducing, and methanogenesis/fermentation media), threemicrocosms were prepared including one with 5% (v/v) of1 M HCl added in order to create abiotic control conditions.Fifty milliliters aliquots were distributed in 100-mL Wheatonserum bottles sealed with butyl rubber stoppers (Bellco Glass,Inc). Benzene, toluene, ethylbenzene, o-, m-, and p-xyleneswere finally added (100 ppm final concentrations; Sigma–Aldrich). All manipulations were done in an anaerobic glovebox (Getinge La Calhene, France) under an atmosphere of95% N2 and 5% H2. Incubations were performed under staticconditions at the deep aquifer in situ temperature of 37◦C in thedark.

Subcultures in sulfate reducing conditions were preparedoutside the glove box using synthetic water (0.5 g.L−1

NH4Cl, 0.1 g.L−1 MgCl2.6H2O, 2 g.L−1 Na2SO4, 0.06 g.L−1

CaCl2,2H2O, 0.5 g.L−1 NaCl, 0.3 g.L−1 KH2PO4, 0.3 g.L−1

K2HPO4, 1 mL.L−1 of an anoxic solution of trace-elements(Eichler and Pfennig, 1986), the composition of which mimickedthat of the formation water. Culture media were sterilized byautoclaving for 20 min at 120◦C and immediately flushed undera stream of O2-free N2 gas and cooled to room temperature priorto the addition of sterile and anoxic solutions of 1 mL.L−1ofvitamins, 2 g.L−1 FeCl2.4H2O, and 2.55 g.L−1 Na2S.H2O. Themedium was adjusted to pH 8 and 50 mL aliquots weredistributed in Wheaton serum bottles sealed with butyl rubberstoppers under a stream of O2-free N2. A 10% inoculum andBTEX (100 ppm) were finally added. Chronology of the differentexperiments during this study is provided in Figure 2.

Analytical ProcedureAqueous samples (0.3 mL) were collected by syringe throughthe stoppers, transferred to chromatographic vials and acidified(10 μL of 3 N HCl) for monitoring BTEX degradationperiodically by SPME/GC/FID with an autosampler CombiPal (CTC Analytics) coupled with a gas chromatograph7890A (Agilent Technologies) equipped with a flame ionizationdetector. BTEX was absorbed in headspace vials during10 s with a micro-extraction fiber (SPME, Supelco 75 μmcarboxen-PDMS). Desorption time inside the GC injector was100 s. Compounds were separated through an Optima Wax(Macherey–Nagel) column (30 m × 0.32 mm × 0.50 μm).Helium was used as a carrier gas with a constant flowrate of 1 mL min−1. Results were processed as the residualpercentage of BTEX as (Ct/Cc) × 100, where Ct is thehydrocarbon concentration in the microcosm, and Cc thehydrocarbon concentration in the abiotic control microcosm.

The resulting values were normalized to o-xylene as an internalstandard.

DNA Extraction, PCR and ssu rRNAClone Libraries for TaxonomicAssignmentFrom the fifth to eighth subcultures, genomic DNA wasextracted in duplicate from the microbial communities usingthe Powersoil DNA isolation kit (MoBio Laboratories, Carlsbad,CA, USA). For the construction of 16S bacterial rRNA genelibraries, targeted genes were amplified using the PCR CoreKit Plus (Roche Diagnostics) with the primer sets 8F/1492Ror 8F/B926R (Lane, 1991; Weisburg et al., 1991). DNAamplicons were purified, cloned, sequenced, and analyzed aspreviously described (Stackebrandt and Goebel, 1994; Coleet al., 2003; Berlendis et al., 2010). From the different clonelibraries, 226 clones were randomly sequenced. Phylogeneticanalyses were carried out after aligning related sequences usingthe Muscle program (Edgar, 2004). Ambiguous regions wereremoved using Gblocks (Castresana, 2000) and phylogenetictrees were constructed using the maximum likelihood methodimplemented in the phyML program v 3.0 (Guindon andGascuel, 2003). Reliability for internal branch was assessedusing the aLRT test (Anisimova and Gascuel, 2006). The 16SrRNA gene sequences reported in this study were deposited inGeneBank database with accession No. KR061296 to KR061298.Archaeal primers tested were primer couples A9F (Vetrianiet al., 1999)-U1492R, A9F/A958R (DeLong, 1992), and A109F(Grobkopf et al., 1998)-A958R. Amplification of the genecoding the α-subunit of benzylsuccinate synthase was carriedout using the “semi-nested” protocol described previously forDesulfotomaculum sp. OX39 (Winderl et al., 2007) or withthe primers set 7768F/8543R designed by von Netzer et al.(2013).

Determination of Isotopic Fractionationof 13C-BenzeneFor the determination of isotopic fractionation, the bacterialactivity was stopped in four microcosms on the seventhsubculture at different stages of 13C-benzene (SIGMA)biodegradation by acidification to pH 2 with 15% HCl. Flaskswere conserved at −20◦C until analysis.

Hydrogen isotope analyses were performed in duplicateby SPME-GC-TC-IRMS using a Hewlett Packard 6890gas chromatograph connected to a Delta plus XLTM massspectrometer with a GC/TC interface (Finnigan MAT). Thegas chromatograph was equipped with a DB-PETRO column(100 m × 0.25 mm × 0.5 μm film, J&W Scientific). Heliumwas used as a carrier gas with a flow rate of 1.7 mL.min−1 forhydrogen isotope analysis. The temperature program startedat 35◦C for 20 min isothermally, was increased at a rate of2◦C.min−1 to 315◦C and maintained isothermally during50 min. Vienna Standard Mean Ocean Water (VSMOW) wasused as the standard for the detection of hydrogen isotope ratiosand results were reproducible within ±2.50/00.






Carbon isotope analyses were performed in duplicate bySPME-GC-C-IRMS using the same GC as described previouslyconnected to a Delta plus XLTM mass spectrometer with a GC/Cinterface (Analytical Precision). The column, carrier-gas andtemperature program used were the same as for hydrogen isotopeanalyses. Vienna Pee Dee Belemnite (VPDB) was used as thestandard for the analysis of carbon isotope ratios (Coplen et al.,2006) and results were reproducible within ±0.30/00.

Element isotope ratios “δhE” (where h is atomic number) areexpressed in delta notation in per mil (�) and will be specificallydesigned for hydrogen and carbon, δH and δC, respectively:δhE [0/00] (δH or δC) = [Rsample–Rstandard/Rstandard] ∗ 1000.Rsample and Rstandard are the 13C/12C or 2H/1H ratios of thesample and of the internal standard, respectively. Enrichmentfactors for hydrogen and carbon were determined accordingto the logarithmic form of the Rayleigh equation using deltanotation (�) as previously described (Elsner et al., 2005;Fischer et al., 2008). ln (Rt/R0) = (ε /1000) ∗ ln(Ct/C0) whereRt/R0 = (δhEt + 1000)/(δhE0 + 1000). Rt and Ct are, respectively,the isotopic composition and the concentration of the compoundat a given time t; R0 and C0 the same variable at the startingpoint of the reaction. ε [0/00] represents an isotopic enrichmentfactor calculated from the slope of the plot ln (Ct/C0) versusln (Rt/R0) multiplied by 1000 giving the ε-value in per mil.The factor �bulk expresses the slope of the linear regression forcarbon and hydrogen discrimination: ∧bulk =�δ2 Hbulk/�δ13

Cbulk. In agreement with previous studies, Ct/C0, which is theresidual concentration of the compound at the time t, is calledf and B [%] is a parameter expressing the extent of the benzenebiodegradation such as B [%] = (1–f ) ∗100.

Enrichment factors were then corrected by consideringenrichment factors specific for the reactive position(ε reactive position). As benzene is a symmetrical moleculewith potentially six reactive carbons and six hydrogen atoms(Fischer et al., 2008), the AKIE (Apparent Kinetic Isotope Effect)considering the intramolecular competition of carbon andhydrogen atoms, is calculated according the following equation(Elsner et al., 2005): AKIE = 1/(1 + z ∗ ε reactive position/1000)with z = 6 (number of atoms of an element in identical reactivepositions).

Microscopic Observation and CellCountsThroughout the study, microcosms were sampled and observedunder phase contrast microscopy. Total cell counts wereperformed by DAPI-staining (4′,6′-diamidino-2-phenylindole,Sigma–Aldrich) with an Olympus BX60 epifluorescencemicroscope equipped with a monochrome camera (12 bits,QIClick) and with a mercury light source. Formation water(18 mL) was fixed on-site with 2 mL of 10% borax-bufferedformaldehyde (37%, Sigma–Aldrich) and stored at 4◦C. Tenmilliliters of sample were stained with 0.5 mL DAPI stocksolution (200 μg.mL−1) then filtered onto 0.2 μm pore-sizeblack polycarbonate filters (Millipore) under vacuum. Forthe eighth subculture, counterstaining was done by filtering amixture of 22 μL of culture with 1 μL of DAPI stock solution

onto 0.2 μm pore-size black polycarbonate filters (Millipore)under vacuum. For each filter, 10 randomly selected fields werecounted.

Inhibition TestsOn the eighth subcultures, BTE degradation inhibition testswere carried out by monitoring the biodegradation periodicallyand injecting sodium molybdate, Na2MoO4 (10 mM finalconcentration) and sodium 2-bromoethanesulfonate or BES,BrCH2CH2SO3Na (2 mM final concentration) through butylstoppers just after the beginning of the biodegradation.

RESULTS

Benzene, Ethylbenzene, and TolueneRemoval Kinetics Along SuccessiveEnrichmentsDeep aquifer water from the original microcosm (November2000) showed degradation of ethylbenzene after 900 daysincubation under sulfate-reducing conditions. The firstsubculture (February 2002) was able to degrade ethylbenzene in100 days and showed the beginning of toluene degradationwhereas the second subculture (July 2002) degradedethylbenzene in 100 days, toluene in 150 days and benzenein 270 days (Figures 1A and 2). The ability to sequentiallydegrade ethylbenzene, toluene, and benzene (BTE) was observedfor the next enrichments under sulfate-reducing conditions(from the second subcultures to the eighth ones), but no removalof xylene isomers was observed (Figures 1 and 2). Degradationwas not observed in the abiotic controls, although there was slowmono-aromatic hydrocarbon absorption by the butyl septa asreported in similar studies (Shen and Sewell, 2005; Holmes et al.,2011). After 38 months of incubation, no BTEX removal wasdetected in the presence of the electron acceptors: nitrate, ironIII, and CO2. Eight successive enrichments were achieved undersulfate-reducing conditions during the eight following years.The initial observation of the biomass in the formation watercollected in November 2000 and before any transfer, showeda low bacterial biomass (8.5 × 103 cells.mL−1) with apparentlow morphological cell diversity. Subcultures of the BTE-degrading microcosms under sulfate-reducing conditions lead toa significant gain in biomass along the successive enrichmentsobtained years after years (Figure 1B). Degradation started aftera lag phase lasting from 50 to 120 days and benzene degradationwas complete approximately 4 months later. Whereas BES, aninhibitor of methanogenesis, addition did not show any effect,BTE degradation was significantly stopped as soon as sodiummolybdate (NaMoO4), a specific inhibitor of sulfate reduction,was injected as shown in Figure 1C. It was particularly apparentwith ethylbenzene where the removal process was stopped at60% immediately molybdate was introduced. No further BTEdisappearance was then observed once MoO4 was introduced.We also tested the BTE-degrading microbial community withethylbenzene (100 ppm), or toluene (100 ppm), or benzene(100 ppm) as the sole carbon and energy sources. In these






FIGURE 1 | (A) Degradation of benzene, toluene, and ethylbenzene (BTE) during the second subculture (07/2002); Filled squares : ethylbenzene, filled diamonds :toluene, filled triangles : benzene, cross : o-xylene as internal standard. (B) Degradation of BTE along increase of observable biomass during the eighth subculture(04/2008); Filled squares : ethylbenzene, filled diamonds : toluene, filled triangles : benzene, cross : o-xylene as internal standard and filled circles: cells.mL−1.(C) Effects of inhibitors addition on BTE biodegradation during the eighth subculture (04/2008); Arrow indicates the addition at day 139 of sodium molybdate (MoO4,10 mM) or bromoethanesulfonate (BES, 2 mM); Filled squares : ethylbenzene + BES, filled diamonds : toluene + BES, filled triangles : benzene +BES, cross :o-xylene as internal standard + BES or + MoO4, open squares : ethylbenzene + MoO4, open diamonds : toluene + MoO4, open triangles : benzene + MoO4. Startlevels of BTEX were 100 ppm.






FIGURE 2 | (A) Degradation of benzene, toluene, and ethylbenzene (BTE) during the original microcosm (10/2000) incubation; (B) Degradation of BTE during thefirst subcultures (02/2002); (C) Degradation of BTE during the second subcultures (07/2002); (D) Degradation of BTE during the fifth subcultures (01/2005).(E) Degradation of ethylbenzene, or toluene, or benzene during the sixth subcultures (04/2008). Filled squares: ethylbenzene, filled diamonds: toluene, filled triangles:benzene, cross: o-xylene as internal standard. Start levels of BTEX were 100 ppm.






last assays, the toluene and the benzene removal rates werehigher than in the BTEX mixture enrichment with a completedisappearance within 3 months (Figure 2).

Isotopic Fractionation Associated toAnaerobic Benzene DegradationFurther investigation performed in the benzene-only amendedmicrocosm showed that organic chemistry of the residualbenzene was also affected along the degradation (noted B [%]).The initial δ13C and δ2H values of the labeled benzene were−25.2 ± 0.1� and −43.5 ± 0.7�, respectively. The analysisof the residual benzene fraction revealed a significant andregular increase of δ13C up to −21.2 ± 0.1� and δ2H upto 58.0 ± 1.4� linked to the extent of benzene degradation(Figure 3). Sterile controls under sulfate-reducing conditionswith different benzene concentrations added showed no isotopicfractionation for carbon and hydrogen, with stable carbon andhydrogen isotopes signatures over time (data not shown). Thespecific apparent kinetic effect of the isotopic fractionation forcarbon and hydrogen (AKIEC and AKIEH) along the benzeneremoval were, respectively, 1.0146 ± 0.0009 and 1.5184 ± 0.0283and were derived from enrichment factors (εC and εH) given inTable 1; Figures 4A,B and 5. The general combined effect ofcarbon and hydrogen isotopic fractionations was gaged by theratio ∧ (∧ = 23.8 ± 0.4) obtained by the dual plot analysis ofthe carbon vs. hydrogen isotope fractionation range along thedegradation (Figure 4C).

Microbial Characterization of theHydrocarbonoclastic EnrichmentCell counts in the BTE-degrading enrichments showed that BTEremoval was linked to a fourfold increase of cells in microcosmsincreasing from 3.6 × 106 to 1.2 × 107 cells.mL−1 (Figure 1B).However, the shortest doubling time about approximately 20 dayswas extremely low. Spores were observed from the initialenrichments (November 2000) and were still observed alongthe enrichments. No evidence of archaeal populations could be

FIGURE 3 | δ13C and δ2H of residual benzene fraction versus benzenedegradation rate under sulfate-reduction. Filled circle: δ13C; clear square:δ2H. Benzene degradation refers to B [%], see Experimental proceduressection. Start level of benzene was 12 ppm.

detected by biomolecular approaches (Archaeal 16S rRNA geneamplification). The clone libraries analyses based on 8F-926Ror 8F-1492R amplicons revealed only two distinct phylotypes,both affiliated to the genus Desulfotomaculum (Figure 6). Thecomparison of the two related nucleic sequences based on1492 nucleotides exhibited divergence above 5.7% betweenthese two phylotypes. The phylotype Bc107 covered 98% ofthe clone libraries from the both clone libraries obtained byboth primers couple, the second one, so-called, Bc105 clusteredonly 2% of the total clone library. Complementary microscopicobservations and biomolecular approaches by t-RFLP analysisconfirmed the low diversity obtained with only two distinctpeaks (data not shown). The dominant phylotype Bc107shared the closest affiliation with environmental sequencesobtained from borehole water in a deep South African goldmine (clone TTMF126, accession number AY741686). Themore closely related environmental sequences for the minorphylotype Bc105 were derived from a 896 m-deep aquiferlinked to the South African gold mine (96% similarity withthe clone DR9IPCB16SCT7, accession number AY604051) and apetroleum-contaminated soil (96% similarity with the clone EKCK572, accession number JN038217). Bc107 population sharedthe closest sequence similarity with thermophilic and moderatethermophilic strains in the cluster Ia of the Desulfotomaculumgenus (Gram-positive Bacteria), such as D. putei isolated fromthe deep subterranean biosphere at 2.7 km depth (Liu et al.,1997), D. hydrothermale isolated from a terrestrial hot spring(Haouari et al., 2008), or D. varum a moderately thermophilicbacterium from a 66◦C- Great Artesian Basin (Ogg and Patel,2011).

DISCUSSION

Assessment of Biodegradation Processat the Origin of the BTE-and BenzeneRemoval Under Sulfate-ReducingConditionThe natural gas stored in underground gas storage aquifers ismainly composed of methane but also contains traces of othercompounds, which include BTEX, with concentrations in partsper billion (ppb). The majority of BTEX is withdrawn at thesame time as the natural gas is extracted from the undergroundreservoir; however, during storage a part of the BTEX dissolvesin the formation water where these compounds are undesirable.In surface environments, or in shallow aquifers with directinfluences from surface environments, microorganisms areexposed to hydrocarbons naturally present in the environment(alcanes, terpenoïds) or introduced by human activities (oilspill). In the case of very deep environments (below −100 m),ecosystems have remained remarkably stable over geologicaltime. After 150 million years isolated from the surface, gasinjection with the input of a trace amount of organic matterrepresents an unknown stress to indigenous microorganismswhich could lead to the selection of specific populations amongthe indigenous microbial community.






TABLE 1 | Carbon and hydrogen isotope enrichment factors retrieved in this study and compared with literature data obtained during anaerobicdegradation of benzene.

Initial benzeneadded [μM]

εC bulk [�] ±95 % CI [�]

R2 εH bulk [�] ±95 % CI [�]

R2 Reference

Pure culture

Ralstonia picketti VKOl(omc) 885 −1.7 ± 0.2 0.98 −11 ± 4 0.86 Fischer et al., 2008

Cupriavidus necator ATCC 17697 (oxic) 1180 −4.3 ± 0.4 0.99 −17 ± 11 0.89 Fischer et al., 2008

Burkholderia sp. (oxic) 700 −3.5 ± 0.3 0.97 −11 ± 2 0.91 Hunkeler et al., 2001

Acinetobacter sp. (oxic) 700 −1.5 ± 0.8 0.99 −13 ± 1 0.99 Hunkeler et al., 2001

Azoarcus denitrificans strain BC (oxic) 603 −2.6 ± 0.8 0.97 −16 ± 4 0.97 Fischer et al., 2008

A. denitrifans strain BC (Chlorate-reducing) 462 −1.5 ± 0.5 0.86 −28 ± 6 0.98 Fischer et al., 2008

Mixed cultures

Nitrate-reducing, mixed negative 250 −2.2 ± 0.4 0.98 −35 ± 6 0.91 Mancini et al., 2003

Sulfate-reducing, mixed negative 192 −3.6 ± 0.3 0.92 −79 ± 4 0.79 Mancini et al., 2003

Sulfate-reducing, mixed negative 192 −1.9 ± 0.3 0.97 −59 ± 10 0.99 Fischer et al., 2008

Methanogenic, mixed negative 750 −1.9 ± 0.1 0.98 −60 ± 3 0.92 Mancini et al., 2003



Sulfate-reducing, enriched positive 450 −2.5 ± 0.2 0.97 −55 ± 4 0.93 Bergmann et al., 2011

Iron-reducing, enriched positive 200 −3.0 ± 0.5 0.93 −56 ± 8 0.93 Bergmann et al., 2011

Sulfate−reducing, enriched positive 400 −2.4 ± 0.3 0.91 −57 ± 0.0 0.98 This study

FIGURE 4 | (A,B) Double logarithmic plot according to the Rayleigh equation expressing changes in isotopic composition and compounds concentrationalong time for carbon and hydrogen during anaerobic degradation of benzene; (C) Dual isotope plots of �δ2H versus �δ13C for anaerobic benzenebiodegradation giving the � values as the slope of the regression. Dashed lines in all graphes represent the corresponding 95% confidence intervalsfrom duplicate analysis.

Our results showed that a deep subsurface confined pristineaquifer can hold microbial communities acting as a keyplayer in the natural attenuation of benzene and alkylbenzenes.

Our evidence is consistent with a biological, rather thana physical–chemical process, causing the disappearance ofbenzene, toluene and ethylbenzene (BTE): (i) repeatability






FIGURE 5 | Biplot of AKIEC vs. AKIEH values from data retrieved in this study and in literature dedicaced to benzene biodegradation. Redox conditionare deduced from the culture condition where the biodegradation was observed. 1: methanogenic consortium (Mancini et al., 2008); 2: methanogenic consortium(Mancini et al., 2008); 3: sulfate-reducing consortium (Fischer et al., 2008); 4: sulfate-reducing consortium, this study; 5: sulfate-reducing consortium (Bergmannet al., 2011); 6: iron-reducing consortium (Bergmann et al., 2011); 7: nitrate-reducing consortium (Mancini et al., 2003); 8: Nitrate-reducing consortium (Manciniet al., 2008); 9: nitrate-reducing consortium (Mancini et al., 2008);10: nitrate-reducing consortium (Mancini et al., 2003); 11: Ralstonia Pickettii (Fischer et al., 2008);12: Azoarcus denitriificans (Fischer et al., 2008); 13: Burkholderia sp. (Hunkeler et al., 2001); 14: Cupriavidus necator ATCC 17697 (Fischer et al., 2008); 15:Acinetobacter sp. (Hunkeler et al., 2001).

over time (eight subcultures); (ii) degradation of benzene,or toluene, or ethylbenzene as sole carbon sources but noxylene degradation; (iii) an increase of biomass linked to thedegradation of the monoaromatic hydrocarbons, in particularthe ethylbenzene (Figures 1 and 2). Isotopic fractionationstudies along with biological processes have been reported fordecades as a promising technique for in situ direct evidencesof biodegradations (Borden et al., 1995; Kelley et al., 1997;Ahad et al., 2000; Meckenstock et al., 2004; Zwank et al., 2004;Bergmann et al., 2011; Braeckevelt et al., 2012). Here, the extentof carbon isotope fractionation was about 4� which is inagreement with the extent of isotopic fractionation expectedfor this compound for biodegradation (>2�); for hydrogenisotope fractionation, the change was about 100� which isin the same range as reported in the review by Braeckeveltet al. (2012) with values >20� as a proof of biodegradationprocess.

Other evidence, strongly suggests that sulfate-reducingmicroorganisms are involved in the BTE degradation. Suchas the lack of degradation with electron acceptors otherthan sulfate (nitrate, iron(III), carbon dioxide). Archeal 16SrRNA gene was not amplified suggesting the absence ofthis domain in this community, in particular methanogenicarchaea. This was confirmed by the inhibition of degradationwith molybdate but not with BES. Additionally, the dual

plot analysis of CSIA with C and H showed that our ∧value (23.8 ± 0.4 with R2 = 0.98; Figure 4C) was wellintegrated with literature values obtained in low redox potentialconditions including fermenting, methanogenic and sulfate-reducing benzene-degrading enrichments (∧ = 22–28). Incontrast, all these values strongly differ with ∧ reported inliterature for benzene degradation in higher redox conditionslike in nitrate-reducing condition (∧ = 12–16) (Mancini et al.,2003, 2008; Fischer et al., 2007, 2008; Bergmann et al., 2011;Gieg et al., 2014). Moreover, enrichment factors obtainedduring anaerobic benzene degradation under sulfate-reducingconditions showed the influence of microbial composition on theextent of the isotopic fractionation of C and H among sulfate-reducing benzene degrading enrichments: our results agree withvalues obtained by Bergmann et al. (2011) with a Gram-positiveenriched benzene degrading community (εC = −2.5± 0.2�withR2 = 0.97; εH = −55 ± 4� with R2 = 0.93) but are differentfrom values obtained with mixedGram-negative sulfate-reducingenrichments (εC = −3.6± 0.3�with R2 = 0.92; εH = −79± 4�with R2 = 0.79 [Mancini et al., 2003) and εC = −1.9 ± 0.3�with R2 = 0.97; εH = −59 ± 10� with R2 = 0.99 (Fischeret al., 2008)]. Our unsuccessful attempt to isolate a pure strainable to degrade at least one of the BTE compounds could bedue to the existence of an obligatory syntrophism between thetwo detected populations at the origin of the BTE degradation






FIGURE 6 | Maximum-likelihood tree based on 16s rRNA gene (1115 bases) showing the phylogenetic relationship between the both sequencesdetected in microcosms (clone Bc105: minor phylotype and clone Bc107: dominant phylotype) and closest relatives. Reliability values (aLRT values)greater than 50% are given at nodes.

as it has been previously described for the anaerobic degradationof toluene by syntrophic fermentative oxidation with a co-culture containing a sulfate-reducing bacterium (Meckenstock,1999).

The addition of fumarate has previously been shown to bethe mechanism of activation of toluene and ethylbenzene undersulfate-reducing conditions (Rabus and Heider, 1998; Kniemeyeret al., 2003). These reactions are catalyzed by fumarate-addingenzymes (FAEs) which encompass benzylsuccinate-synthase(Bss) and alkylsuccinate-synthase (Ass). PCR amplificationstargeting bssA gene were unsuccessful using previously describedspecific primers (Winderl et al., 2007; von Netzer et al., 2013).The absence of known metabolic genes has also been reportedin other BTEX degrading consortia dominated by Gram-positivebacteria (Hermann et al., 2008; Abu Laban et al., 2009; Bergmannet al., 2011). For two axenic BTEX degrading nitrate-reducingstrains: Dechloromonas aromatica RCB and Dechloromonas sp. JJ

(Coates et al., 2001; Chakraborty et al., 2005), functional genesresearch investigations and genomic analyses did not highlightany known pathways for anaerobic degradation of aromaticssuch as the central benzoyl-CoA pathway for monoaromaticsand benzylsuccinate synthase (bssABC) genes for toluene andm-xylene degradation (Salinero et al., 2009). In our study, theabsence of benzylsuccinate-synthase activity remains unclear.It can be postulated that if present in our community, thisenzyme and its genes are different enough from those previouslydescribed to prevent gene amplification by known primers.Similar difficulties have already been reported for the bssAgene in toluene-degrading communities (Sun et al., 2013) andwith the toluene-degrading strain Desulfotomaculum sp. Ox39(Winderl et al., 2007). Alternatively, the isotopic signatureof carbon and hydrogen obtained both here and previouslyby Bergmann et al. (2011) could be related to an unknownpathway for the anaerobic biodegradation of monoaromatic






compounds and possibly specifically to a lineage among sulfate-reducing Gram-positive biodegraders. Derived values fromisotopic fractionation of carbon and hydrogen, especially AKIECand AKIEH, published here in Figure 5 (1.0146 ± 0.0009and 1.5184 ± 0.0283, respectively) were perfectly integratedwith other AKIE indices available in literature and showed aspecific clustering among other benzene-degrading conditionswith sulfate-respiring cultures. Our data will reinforce the limitedexisting research in term of anaerobic biodegradation of benzeneavailable in the literature (Mancini et al., 2003; Fischer et al.,2008; Bergmann et al., 2011). Dual plot carbon specific isotopeanalysis, especially AKIE indices, initially suggested by Elsneret al. (2005) and reviewed by Braeckevelt et al. (2012) enables awider range of comparisons across the biodegradation kinetics ofdifferent hydrocarbons and remains one of the more promisingmonitoring techniques which may, in the long term, discriminatethe biochemical pathway and energy sources involved during insitu natural attenuation in subterranean environments.

A New Hydrocarbonoclastic MicrobialPopulation Representative of the DeepSubsurface Confined AquiferThe regular observation of spores along the successive BTE-degrading enrichments and benzene-degrading enrichmentsstrongly suggest that spore-forming microorganisms play animportant role in the BTE degradation. Molecular biologyapproaches revealed the presence of only two differentspecies and both belonging to Desulfotomaculum genus(Firmicutes/Clostridia/Clostridiales/Peptococcaceae, Widdel,2006). Desulfotomaculum sp. and related Gram-positive sulfate-reducing bacteria such as Desulfosporosinus sp. are frequentlyencountered in deep environments (Daumas et al., 1988; Nilsenet al., 1996; Ehinger et al., 2009; Aüllo et al., 2013) and aresometimes the main representatives of these communities (Bakeret al., 2003; Moser et al., 2003; Detmers et al., 2004; Moseret al., 2005), especially in several oilfields over the world withgenerally a positive correlation of cell abundance with increasingtemperature with depth (Aüllo et al., 2013; Guan et al., 2013).Their presence in these environments could be linked to theirability to sporulate allowing them to withstand adverse periodsduring burial such as lack of nutrients or heat (O’Sullivanet al., 2015), and their metabolic versatility as demonstrated bytheir ability to respire sulfate, thiosulfate, sulfur, sulfite, metals,and metalloids. Available data on microbial diversity in deepaquifers used as natural gas storage are scarce. Nevertheless,others studies on other deep aquifers revealed the presence ofthis genus sometimes abundantly (Basso et al., 2009; Ehingeret al., 2009; Berlendis et al., 2010). Although members ofDesulfotomaculum can also be found in surface ecosystems, bothpopulations identified in this study are close to environmentalsequences detected in deep subsurface environments supportingtheir ecological relevance (Moser et al., 2005; Gihring et al.,2006; Baito et al., 2015) (AY604051, AY741686, AB910321).On the basis of their sequences encoding the 16S rRNA gene,we assume that we are dealing with two different species.Some members of Desulfotomaculum and Desulfosporosinus

have the ability to degrade hydrocarbons, in particular mono-aromatic hydrocarbons such as toluene, m-xylene and o-xylene(Robertson et al., 2000; Liu et al., 2004; Morasch et al., 2004;Abu Laban et al., 2015). Until now, bacterial isolates have shownthe ability to degrade benzene either in iron-reduction (Holmeset al., 2011; Zhang et al., 2012) or in nitrate-reduction (Coateset al., 2001; Chakraborty et al., 2005; Kasai et al., 2007). Here,we demonstrate that some members of Desulfotomaculumare also able to degrade benzene and ethylbenzene undersulfate-reducing conditions. Despite multiple assays, no pureisolates able to degrade benzene were obtained confirmingthe difficulty to obtain a pure sulfate-reducing strain ableto degrade anaerobically benzene (Vogt et al., 2011; van derZaan et al., 2012), which requires us to hypothesize a possiblesynergy between these two populations of Desulfotomaculum.However, why have these organisms the capacity to degradeBTE in a poorly carbonated sandstone deep aquifer? What weknow about this type of aquifers implies that the indigenousmicrobial communities before gas storage had to be essentiallybased on using CO2 and H2 on the principle of a subsurfacelithoautoautrophic microbial ecosystem (SLiME) as mentionedby Stevens and McKinley (1995) and Basso et al. (2009). Havinga hydrocarbon biodegradation ability could provide a benefitto Desulfotomaculum which can switch to various energysources along with post-diagenetic environmental changes andconsume this type of molecule potentially present as residuesin a fossil organic matter entrapped in rocks. Alternatively,these microorganisms may have the same origin as the injectedgas and would therefore be derived from an oil reservoir. Thishypothesis would explain the ability of these organisms todegrade hydrocarbons, in particular BTEX. The natural gastreatment process after extraction from the oil reservoir, andespecially the dehydration steps would allow only few sporesto resist. Recently, it has been shown that Desulfotomaculumspores could resist triple autoclaving processes (O’Sullivanet al., 2015). Spores could then be transported thousands ofkilometers in pipelines and co-injected with natural gas into deepaquifers. A previous report of the isolation of Desufotomaculumthermocisternum from the North sea oil reservoir able togrow syntrophically with a methanogen (Nilsen et al., 1996),the presence of Desulfotomaculum and methanogen speciesdominating the microbial diversity of a deep gold mine at4–6 km of depth (Moser et al., 2005) support the hypothesis of aversatile metabolism in various subsurface habitats.

Many studies suggest a possible key role of Gram-positiveof members of Clostridia (Winderl et al., 2010) and the familyPeptococcaceae in BTEX biodegradation (Phelps et al., 1998;Da Silva and Alvarez, 2007; Kunapuli et al., 2007; Kleinsteuberet al., 2008; Musat and Widdel, 2008; Oka et al., 2008; AbuLaban et al., 2009; Berlendis et al., 2010; Taubert et al., 2012;van der Zaan et al., 2012; Kuppardt et al., 2014). This studydemonstrates the key role of sulfate-reduction in this community(Figure 1C) and we could deduce that Desulfotomaculum fromthe community (Bc105 or Bc107 or both) degrade the aromatichydrocarbons by direct oxidation. This BTE degradation wouldbe similar to those described by Abu Laban et al. (2009). Theseauthors clustered the highly abundant sequences between the






genera Desulfotomaculum and Pelotomaculum and postulatedthese organisms were key players in benzene degradation withsulfate as the electron acceptor. The slow kinetic removalof monoaromatic hydrocarbons, and benzene in particular,observed in our study compared to kinetics reported in literature(Oka et al., 2008; Abu Laban et al., 2009; van der Zaan et al.,2012) could be linked to (i) an inhibition of anaerobic benzenedegradation by co-contaminants (Edwards and Grbic-Galic,1992; Cunningham et al., 2001; Ruiz-Aguilar et al., 2003; Da Silvaand Alvarez, 2007; Vogt et al., 2011); (ii) microbial metabolismunder extreme energy limitation (Hoehler and Jørgensen, 2013)and (iii) syntrophic consortia requiring optimal conditions(Vogt et al., 2011). Syntrophism between a fermentative anda by-products utilizer could be a logical adaptive strategy ofindigenous lithoautotrophic communities to the presence ofrecalcitrant hydrocarbons. This is supported by a previousinvestigation using DNA-SIP techniques from a sulfate-reducingcommunity degrading benzene enriched from contaminatedgroundwater, showed the likely dominance of still undescribedGram-positive species seemingly active at the early stage ofthe benzene degradation followed by an undescribed Epsilon-proteobacteria assumed to be a hydrogen scavenger (Hermannet al., 2008). In the same way, a study by protein-SIP applied on abenzene-degrading microbial consortium from a shallow aquifer,Taubert et al. (2012) hypothesized the key role of Clostridiales,in particular Desulfotomaculum and Pelotomaculum genera, forbenzene degradation. These authors and others (Kleinsteuberet al., 2008) suggested Peptococcaceae could putatively fermentbenzene and excrete by-products such as acetate and hydrogenwhich would be used by the whole community, in particularDelta-proteobacteria (sulfate-reducers). If this hypothesis is true,the inhibition of sulfate-respiration after adding molybdate couldstop the benzene degradation since the reaction would bethermodynamically unfavorable.

New Bio-Indicator Parameters (AKIEValues, Phylotypes) for BenzeneBiodegradation: Field Applicability inDeep Subterranean EnvironmentsThe selection of an active benzene and alkylbenzenes-degradingcommunity shows that these hydrocarbonoclastic populationscould be persistent and certainly able to sustain in situbiodegradation as long as a substrate is available. Deep subsurfaceconfined aquifers despite restricted access remain key locationpoints for subsurface engineering activity (water resources,oil, and gas industries, geothermal energy, bioremediation,fundamental research interests). However, field biodegradationstudies are currently mandatory for bioremediation, oil recoveryor geological exploration. Indeed, in the context of hydrocarbonbiodegradation strongly linked to aquifers properties (Warrenet al., 2004), petroleum industries, bioremediation experts, andenvironmental ecologists point out the need to systematicallycollect and share various technical information requiring astrong fundamental research background such as microbialecology, isotopic studies, and biogeochemistry (Scow and Hicks,2005; Declercq et al., 2012; Hubbard et al., 2014). Because

of the higher benzene persistence in anoxic environmentsthan its others alkylated derivatives, we focused our study onthis hydrocarbon. Sequences of both phylotypes showed noclose affiliation with any Desulfotomaculum species detected inbiphenyl or benzene-degrading consortia reported in literature(Abu Laban et al., 2009; Selesi and Meckenstock, 2009). Inaddition, both phylotypes were loosely affiliated to any knownhydrocarbonoclastic microorganisms, such asDesulfotomaculumsp. Ox39 (Morasch et al., 2004). Hence the results presentedhere obtained with this new benzene-degrading enrichmentsignificantly extent the diversity of Gram-positive biodegradersfrom a deep subterranean aquifer (Abu Laban et al., 2009;Bergmann et al., 2011: for the last review about microorganismsinvolved in anaerobic biodegradation of petroleum, see Widdelet al., 2010). As microbial diversity in low-energy environmentsis known to contain only a few cultivated microorganisms, withtheir biochemistry and physiology largely unknown (Hoehler andJørgensen, 2013), the additional information provided by DNA-based microbial identification such as AKIE may enable furtherstudies with similar results to bring new evidences and newinsights about these biodegradation processes. For example, aconceptual model of syntrophic biodegradation of hydrocarbonshas been initially suggested by Head et al. (2010) and recentlysupported again by Gieg et al. (2014) where several species couldbe involved.

Although anaerobic benzene biodegradation has been clearlydemonstrated in various conditions, the genetic pathwaysinvolved are still unclear, despite the finding of a putative genecluster (Abu Laban et al., 2010; Vogt et al., 2011). So far, nogenetic probe targeting functional genes exists to show directevidence of anaerobic benzene biodegradation. Therefore, fieldapplications of compound-specific isotope analysis (CSIA) andcomparison with literature data from well-known biodegradationcases are currently ourmost powerful diagnostic tools. Significantoutputs of this work would concern petroleum companies notonly in the context of bioremediation of deep confined aquifers(Declercq et al., 2012) but also for understanding microbial-induced souring in the oil and gas producing reservoirs (Sunet al., 2005; Wilkes et al., 2008; Gieg et al., 2011).

CONCLUSION

In this study, it was shown for the first time that abacterial community composed of only two Desulfotomaculumpopulations can use toluene, ethylbenzene and benzene assole carbon and energy sources in sulfate-reducing conditions.They constitute the simplest model of anaerobic sulfate-reducing hydrocarbon-degrading anaerobes originating fromthe deep subterranean environments ever described. Whilemany studies have shown Pelotomaculum sp., another genus ofPeptococcaceae, as key-players in BTEX biodegradation, this workhighlights the important role of the genus Desulfotomaculumas significant indigenous populations of subsurface habitats,but also as an important agent in the anaerobic degradationof hydrocarbons. In the deep subterranean biosphere, complexlitho-autotrophic microbial network promotes reactions for






hydrogen interspecies electron transfer. We hypothesize selectionof microbial populations able to conduct syntrophic oxidation oforganic carbon could be a logical adaptive response of originallylitho-autotrophic indigenous microbial communities to thepresence of recalcitrant hydrocarbons. Yet, at a time when theexploitation of shale gas and oil is quickly increasing globally, it isnecessary to know whether these still poorly undescribed deepsubterranean environments have the potential of hydrocarbondegradation, and particularly BTEX. Field monitored naturalattenuation (MNA) approaches for subterranean environmentsshould be favored by updating the existing database grouping theidentified microbial phylotypes and isotopic values in the contextof BTEX biodegradation.

AUTHOR CONTRIBUTIONS

TA, SB, and J-F L designed, performed experiments, analyzeddata and wrote the paper; DD and DD performed isotopic

fractionation experiments; SS-L, BS, and JM performed culturalexperiments; DP and CB critically reviewed this paper; MM andARP designed, analyzed data, supervised the project and wrotethe paper; TA, SB, and J-FL are co-first authors.

FUNDING

STORENGY and TIGF are acknowledged for funding theIPREM-EEM team for this research project.

ACKNOWLEDGMENTS

The GIP team of STORENGY is warmly thanked for hisinvaluable involvement in sampling campaigns on undergroundgas-storage sites. The authors wish to thank Barry Cragg (CardiffUniversity, School of earth and ocean sciences) for improving thismanuscript.

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Conflict of Interest Statement: The authors declare that the research wasconducted in the absence of any commercial or financial relationships that couldbe construed as a potential conflict of interest.

Copyright © 2016 Aüllo, Berlendis, Lascourrèges, Dessort, Duclerc, Saint-Laurent,Schraauwers, Mas, Patriarche, Boesinger, Magot, Ranchou-Peyruse. This is an open-access article distributed under the terms of the Creative Commons AttributionLicense (CC BY). The use, distribution or reproduction in other forums is permitted,provided the original author(s) or licensor are credited and that the originalpublication in this journal is cited, in accordance with accepted academic practice.No use, distribution or reproduction is permitted which does not comply with theseterms.










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